High-Throughput Quantitative RT-PCR in Single and Bulk C. elegans Samples Using Nanofluidic Technology

This paper presents a high-throughput reverse transcription quantitative PCR (RT-qPCR) assay for Caenorhabditis elegans that is fast, robust, and highly sensitive. This protocol obtains precise measurements of gene expression from single worms or from bulk samples. The protocol presented here provides a novel adaptation of existing methods for complementary DNA (cDNA) preparation coupled to a nanofluidic RT-qPCR platform. The first part of this protocol, named ‘Worm-to-CT’, allows cDNA production directly from nematodes without the need for prior mRNA isolation. It increases experimental throughput by allowing the preparation of cDNA from 96 worms in 3.5 h. The second part of the protocol uses existing nanofluidic technology to run high-throughput RT-qPCR on the cDNA. This paper evaluates two different nanofluidic chips: the first runs 96 samples and 96 targets, resulting in 9,216 reactions in approximately 1.5 days of benchwork. The second chip type consists of six 12 x 12 arrays, resulting in 864 reactions. Here, the Worm-to-CT method is demonstrated by quantifying mRNA levels of genes encoding heat shock proteins from single worms and from bulk samples. Provided is an extensive list of primers designed to amplify processed RNA for the majority of coding genes within the C. elegans genome.


Introduction
The optimization of single-cell RNA sequencing and qPCR revealed that transcriptional pulses or bursts can lead to massive variation in the number of RNA molecules per cell 1 . Further, these technologies uncovered substantial cellular heterogeneity previously missed by standard bulk transcriptomic measurements. Depending on the context, some single-cell transcriptional variability is caused by mixed cellular composition of tissues. However, even in isogenic cell populations grown under the same environment there is widespread transcriptional heterogeneity 2,3 . This 'biological variability' is increasingly identified as a ubiquitous property of cellular networks, from bacteria to man. In some cases, it can have phenotypic consequences in development, cancer progression, HIV latency, and response to chemotherapy 4,5 .
The nematode Caenorhabditis elegans is a unique model organism with ideal characteristics for studying the causes and consequences of biological variability between individuals. These nematodes are a simple model organism composed of 959 cells, and their transparent cuticle makes them amenable for in vivo imaging studies 6 . C. elegans is a hermaphroditic species that predominantly reproduces through self-fertilization; this resulted in isogenic laboratory strains. Despite isogenicity and controlled culture conditions, many phenotypes and transcripts are variable across individuals, suggesting that stochastic or microenvironmental differences contribute to heterogeneity across individuals 7,8 . Such variability in gene expression has multiple fitness consequences, including variability in the penetrance of mutations, survival, developmental timing, and fecundity 7,8,9 . Due to these features, single-worm studies provide the unprecedented opportunity to study biological variability in a whole organism.
There is a fundamental need in the field to develop and optimize technologies for accurate detection of transcripts at a single-worm level. New technologies, such as single-worm RNA sequencing 10 , RNA sequencing from isolated tissues 11 , and single-cell sequencing 12 are now available for C. elegans. However, a main challenge remains: when monitoring interindividuality, weakly expressed genes often fall below detectable levels 13 . This is particularly relevant for rare transcripts isolated from small amounts of starting material, as there is a well-established, inverse relationship between mean expression and technical variance, often causing rare transcripts to fall below statistical cutoffs 13 . The optimization of high-throughput multiplexed qPCR technologies has proven useful for mammalian single-cell studies, in particular when studying the expression of rare transcripts 14,15 . This technology can also be used for benchmarking and validation purposes of other single-worm technologies. more reliable than traditional guanidium thiocyanate-phenol-chloroform extraction, because worms are directly added to the lysis buffer, skipping direct isolation of easily degradable RNA. Second, utilizing nanofluidic technologies significantly increases the number of samples and targets that can be run simultaneously. In this paper, two chips are compared: a single-array chip and a multi-array chip. A single-array chip can run 96 single worms and 96 primer sets, resulting in 9,216 reactions per experiment. To accomplish a similar throughput using standard qPCR technologies would require 96 separate qPCR experiments, using 96 well plates. The smaller and more flexible multi-array chip consists of six 12 x 12 arrays resulting in 864 reactions. The method's superior reliability and sensitivity are boosted by nanofluidic technology and by the introduction of a preamplification step. The method presented in this paper is meant to be used together with a state-of-the art statistical algorithm to extract biological variance. This article presents the protocol for rapid cDNA preparation and high-throughput qPCR for both single-worm and batch worm samples; the algorithm will be published elsewhere. For this protocol, the organization of each chip should be prepared prior to the experiment. Table 1 and Table 2 show examples of these plans for a multi-array and single-array chip, respectively. There are also overviews of the Worm-to-CT protocol detailed in Figure 1 and running the multi-array and single-array chips in Figure 2.
Protocol NOTE: Throughout this protocol Caenorhabditis elegans is referred to as "worm" or "worms". A variety of C. elegans strains can be ordered through online databases or by directly contacting labs that use the model organism. Part I of this protocol (sections 1-3) describes cDNA preparation through the Worm-to-CT protocol. Part II of this protocol (sections 4-13) describes running high-throughput RT-qPCR using nanofluidics, adapted from a protocol developed by Fluidigm 16 . This protocol applies to the use of the two types of nanofluidic chips defined earlier, the single-array chip, which can monitor 96 targets into 96 samples (9,216 RT-qPCR reactions total), or the multi-array chip, which functions as subunits of 12 target x 12 samples. Every multi-array chip contains six independent arrays that can be run together or separately. For instance, using a whole multi-array chip can monitor 72 targets x 12 samples (or vice versa), or 36 targets x 24 samples (or vice versa). For further information regarding any of the materials used in this protocol, refer to the Table of Materials.

RT-qPCR primer validation
NOTE: Real-time primers were designed based on the recommended properties originally issued by MIQUE guidelines 17 . To make primers specific for processed RNA, products were designed such that the two primers bound to either side of at least one splice junction. Requirements for suitable primers included 20%-80% guanine and cytosine content, a melting temperature of 58-60 °C, a difference in melting temperature between primer pairs of ≤0.5 °C and a product length of 70-120 bp. The sequence of the primers generated can be found in Supplemental Table 1. Open source code for the scripts used to generate the primers can be found at https://github.com/s-andrews/wormrtpcr. Primer pairs for transcripts with splice sites were designed so that they lie in two exons flanking an intron but were CAUTION: When handling liquid nitrogen wear Cryo-Gloves as well as protective eyewear and adhere to standard clothing regulations, because contact with the skin or eyes can cause serious frostbite injury.

5.
Freeze-thaw the PCR tubes 10x by transferring them between liquid nitrogen and a ~40 °C water bath. Leave the tubes in the liquid nitrogen for a minimum of 5 s to ensure the samples are completely frozen. Leave the tubes in the water bath until the samples thaw. Do not leave in for a longer period, as this leads to RNA degradation.
NOTE: Tubes can be left in liquid nitrogen for an extended period of time, as samples are frozen to approximately -200 °C, reducing RNA degradation. This should not, however, be a protocol stopping point, because liquid nitrogen evaporates rapidly.

6.
Mix the samples on a thermal mixer (

7.
While the samples are being mixed, thaw the stop solution on ice.

8.
Spin the samples down using a tabletop microcentrifuge (

Reverse transcription
NOTE: For reverse transcription of single worms, the results shown here were generated using the reagents provided with the nanofluidic chips (option 2 in Figure 1). The reagents highlighted in option 2 of Figure 1 were also used for reverse transcription of pooled samples. Either method works interchangeably for the different sample types.

1.
Reverse transcription of single worms

1.
In an RNase-free hood, add 1.25 μL of reverse transcription mix (

2.
Take 5 μL of the lysis solution and stop solution mix from step 2.8 and add it to the fresh PCR tube containing the reverse transcription mix.
NOTE: The cDNA produced can be stored at -20 °C before proceeding to amplification and data collection using high-throughput qPCR.

1.
In an RNase-free hood, prepare a master mix consisting of 12.5 μL of 2x RT buffer, 1.25 μL of 20x RT enzyme buffer, and 0.25 μL of nuclease-free water per sample. Add 14 μL of master mix to 11 μL of lysis solution and stop solution mix from step 2.8.
NOTE: When dealing with a large number of samples this can be performed in 96 well plates.

2.
Run the samples through a thermocycler using the following reverse transcription program: 37 °C for 60 min, 95 °C for 5 min, and 4 °C for ∞ .
NOTE: Generally, the products come to a final volume of 25 μL, in which case 75 μL should be added. However, due to condensation the final volume can vary. Therefore, adjust accordingly to make a 1:4 ratio in the final solution. This dilution step does not apply if performing qPCR on single worms. The cDNA produced can be stored at -20 °C before proceeding to amplification and data collection using high-throughput qPCR.

1.
Prepare a forward/reverse (F/R) primer stock for each pair of primers at 50 μM final concentration. Mix the same volume of forward and reverse primers at 100 μM each.

2.
Combine 1 μL of 50 μM F/R primer stock for each primer pair to be tested. Add the DNA suspension buffer up to a total volume of 100 μL.
NOTE: The stock primer concentrations here differ from those described in the manufacturer's protocol 16 but retain the same final concentration of 500 nM.

1.
Prepare a master mix containing 1 μL of preamplification mastermix (Table of Materials), 0.5 μL of the pooled primer mix (step 4.2), and 2.25 μL of nuclease-free water per reaction with a 10% overall surplus volume.

2.
In a 96 well plate, aliquot 3.75 μL of the master mix into as many wells as required for the number of samples to be run.

4.
Cover the plate with 96 well sealing tape, briefly vortex, and centrifuge with a tabletop plate spinner. Transfer to a thermocycler and run the following program: 95 °C for 2 min, 15 cycles of denaturation at 95 °C for 15 s, annealing/extension at 60 °C for 4 min, and 4 °C for ∞ .
NOTE: The manufacturer recommends from 10-20 cycles for the preamplification reaction 16 . This protocol recommends 10 or 15 cycles depending on the expression levels of the target genes.

Exonuclease I treatment
NOTE: This is to remove unincorporated primers from preamplification.

1.
Prepare an exonuclease I mix containing 0.2 μL of exonuclease I reaction buffer (

2.
Remove the 96 well plate (step 5.4) from the thermocycler, centrifuge with the tabletop plate spinner, and carefully remove the seal. Add 2 μL of the exonuclease I mix to each preamplification reaction. Reseal, centrifuge, and place the 96 well plate back into the thermocycler using the following program: 37 °C for 30 min, 80 °C for 15 min, and 4 °C for ∞ .

3.
Take the samples out of the thermocycler and dilute them 1:5 by adding 18 μL of 1x Tris EDTA buffer (Table of Materials).
NOTE: It is possible to keep the cDNA samples at -20 °C for later use. The manufacturer's protocol suggests potential dilutions of 5x, 10x, or 20x at this stage 16 , depending on the expression level of the targets of interest.

Preparing the assay mixes
NOTE: Assay mixes can be prepared in 384 well plates, as the wells have the same spacing as the nanofluidic chips, making loading easier.

1.
Preparing assay mixes for a multi-array chip

1.
Prepare a master mix consisting of 2 μL of 2x assay loading reagent (

2.
Add 0.4 μL of the 50 μM F/R primer stock prepared in step 4.1 to the appropriate wells according to the prepared plan.
NOTE: This provides a total of 4 μL of assay mix per well, with a surplus of 1 μL.

2.
Preparing assay mixes for a single-array chip

1.
Prepare a master mix consisting of 3 μL of 2x assay loading reagent and 2.4 μL of DNA suspension buffer for each well according to the prepared plan. Aliquot 5.4 μL of this master mix per well into a marked 384 well plate.

2.
Add 0.6 μL of the 50 μM F/R primer stock to the appropriate wells according to the prepared plan.
NOTE: This provides a total of 6 μL of assay mix per well, with a surplus of 1 μL.

Preparing the sample mixes
NOTE: Sample mixes can be prepared up to 1 day in advance and stored at 4 °C.

1.
Preparing samples for a multi-array chip

1.
Prepare a sample master mix consisting of 2 μL of 2x fluorescent probe supermix with low ROX ( NOTE: This gives a total of 4 μL, with a surplus of 1 μL.

2.
Preparing samples for a single-array chip

1.
Prepare a sample master mix consisting of 3 μL of 2x fluorescent probe supermix with low ROX (

2.
Pipette 2.7 μL of each preamplified and exonuclease I treated sample from step 6.3 into the appropriate wells according to the prepared plan.
NOTE: This gives a total of 6 μL, with a surplus of 1 μL. If there are any wells to be run without a sample these must be loaded with sample master mix and 2.7 μL of water instead of cDNA. This is recommended for both chip types. The machine needs low ROX in every inlet in order to detect the chip's grid.

Priming the nanofluidic chip
NOTE: A multi-array chip only needs to be primed on the first run. If there are subsequent runs of the same chip this stage can be skipped. These steps are the same for both chip types.

1.
Slowly and carefully, inject the full 150 μL of the control line fluid from the syringes included into the accumulators of the chip. Ensure that no control liquid touches the chip by holding the chip at a 45° angle and holding the tip of the syringe away to avoid spillage.

3.
Place the chip into the nanofluidics PCR priming machine (

1.
Remove the barrier plugs sequentially as loading takes place. This reduces the chance of misloading wells.

2.
Transfer either 3 μL for a multi-array chip or 5 μL for a single-array chip of each primer assay mixes and sample mixes to the corresponding inlets of the nanofluidic chip according to the prepared plan. Make sure not to introduce bubbles, which can cause the total volume transferred to be smaller than desired.
NOTE: If there are any wells that will be run without a primer it is important for these to be loaded with master mix, substituting the primer volume with water. This applies to both chip types. At this stage, it can be easier to place the chip on a dark surface, which will allow the wells to be seen more easily.

Running the nanofluidic chip
NOTE: The first time running a multi-array chip, set up the tracking file by selecting Tools | Flex Six Usage Tracking, click New, enter a file name, and select a location before clicking Done.

1.
Open the data collection software. Click Start New Run. Place the loaded chip into the nanofluidics thermocycler with the barcode facing outwards.

2.
Choose the project setting if applicable, then click Next | Load. If loading a multi-array chip, select the partitions (arrays) to be run.

3.
Select the application Reference Probes, then change the application type to Gene Expression, and change the passive reference to ROX. Select the Single Probe Assay, change the probe type to Eva Green, and click Next.

4.
Select the thermal cycling protocol GE FLEX six Fast PCR+Melt v1 to run a multi-array chip, or the protocol GE 96.96 Fast PCR+Melt v2 to run a single-array chip.

5.
Confirm that Auto Exposure is selected and click Start Run.

Post chip run
NOTE: This section is only necessary for multi-array chips when not using the entire chip.

1.
Take the chip out of the nanofluidics thermocycler and load into the nanofluidics PCR priming machine and run the Post Run (153x) script, which lasts 5 min.

2.
Label the plugs used for personal reference.

1.
Open the data in the 'Real-Time PCR Analysis' software (Table of Materials).
Check the melting peak temperature for every primer pair tested. Eliminate results exhibiting more than one melting temperature peak, for a given primer pair.
NOTE: Multiple peaks only appear occasionally, presumably when primer pairs forms dimers, or from interactions of target primers with other primers in the pooled primer mix.

2.
Export the data as a 'heatmap' spreadsheet file and eliminate failed samples or primers.

3.
Analyze data using the standard Delta-Ct method 21 . For statistical evaluation perform one-way ANOVA on relative expression levels.

Validation of Worm-to-CT as a cDNA preparation method
To test if the Worm-to-CT protocol is a valid cDNA extraction method, it was compared to standard guanidium thiocyanate-phenol-chloroform extraction methods. The results are shown in Figure 3, where cDNA was prepared from an average of ~1,000 worms using standard guanidium thiocyanate-phenol-chloroform extraction techniques 22 and from 30 worms using the Worm-to-CT method. The samples were heat shocked simultaneously (30 min at 34 °C). Globally, hsp-70 mRNA expression levels per 100 ng of total RNA were comparable using both methods. However, in the case of highest hsp-70 expression (i.e., in N2 following heat shock) expression levels were higher with the Worm-to-CT method, indicating improved sensitivity.
To determine if an expected decrease in hsp expression in hsf-1(sy441) 23 , a mutation in the main transcriptional regulator of molecular chaperones 23,24 , could be reproduced, transcriptional chaperone induction following a brief heat shock was compared. With both methods a decrease in hsp-70 induction was detected in hsf-1(sy441) animals. This was expected, because mutant hsf-1(sy441) animals exhibit a decreased ability to induce chaperones due to a truncation in the transactivation domain of HSF-1. For guanidium thiocyanate-phenol-chloroform extraction hsp70 decreased by 82.7% compared to controls and 92.3% for Worm-to-CT compared to wild type animals ( Figure 3). The results were comparable between both methods and comparable to previous reports 23 . These results indicate that the Worm-to-CT method is a valid alternative to standard cDNA synthesis techniques.

Validation of the nanofluidics PCR platform used to amplify mRNA targets
To test the consistency of the results using nanofluidic qPCR for transcript amplification, the PCR results obtained from the Worm-to-CT bulk method were compared on both a standard qPCR system (Table of Materials) and a nanofluidic qPCR system using a multi-array chip. The fold change in the expression of three different genes, sma-3 ( Figure  4A), sma-10 ( Figure 4B), and dnj-26, was monitored ( Figure 4C) in animals carrying a null allele in dbl-1 (dbl-1(nk3)) 25  respectively. Altogether, these results show that the Worm-to-CT method applied to bulk samples is an efficient and rapid way to extract RNA from few worms and provides reliable data when coupled with either standard PCR systems or high-throughput nanofluidics-based qPCR platforms.

Comparison between the expression levels obtained by bulk samples with averages obtained from single worms
The relative expression levels were calculated using either cDNA obtained from bulk samples (25 worms) or from an average of 36 single worm samples ( Figure 5). Both cDNAs were obtained using the Worm-to-CT method and amplified using nanofluidics PCR technology. As observed in Figure 5A-C, for all chaperones tested (i.e., hsp16.1, F44E5.4, hsp-70), the methods detected comparable expression levels. These results indicate that parameters obtained from single worms are reliable.

Application of Worm-to-CT coupled to nanofluidics technology to estimate single-worm gene expression parameters
Because the single-array chip allows monitoring of up to 96 target transcripts on 96 individual samples, it is therefore well-suited to monitor individual variability in transcript expression between single worms. Figure 6A presents a representative result showing the mean expression of multiple hsp transcripts from single worms following a short heat shock. As observed in the figure, the variability in the expression of transcripts differed dramatically across different genes ( Figure 6A). To gain further insight, the coefficient of variation (CV) was calculated by dividing the standard deviation by the mean of the expression levels 26 ( Figure 6B). Three genes whose CV values have been previously estimated by alternative methods were monitored (unpublished data). Two stable transcripts (ife-1 and Y45F10D.4) and one variable (nlp-29 27 ) showed their expected variability. The graph also clearly depicts the well-known inverse relationship between variability values and expression levels 26 ( Figure 6B).
Technical replicates are of paramount importance to ensure reproducibility when using bulk samples. However, this is not necessarily the case for single-cell experiments 14,15,28 . To determine if the use of technical replicates is necessary for parameter estimation when using single-worm samples, 28 individual worms were harvested, following a short heat shock, and processed using technical triplicates. The CV values calculated from single-worm data obtained in triplicate (blue dots in Figure 7, technical CV) versus those for every transcript obtained from individual worms (red dots in Figure 7, biological variability) were compared. For every transcript tested, the technical CVs were lower than the biological CVs, indicating that technical triplicates were not required for parameter estimation. The fact that technical replicates are not required increases the throughput of the experiment without compromising quality.

Discussion
In this paper, the Worm-to-CT protocol is shown to be a rapid and efficient method to extract RNA from single worms or a small pool of worms. The high-throughput offered by the nanofluidic system makes it ideal for quantification of inter-individual variability measurements. Furthermore, the high sensitivity of this method allows the detection of genes expressed at low levels that fall below detection when using single-worm RNA-seq technologies 9 .
When considering the choice of method to prepare cDNA from single worms. Ly et al. 29 optimized a protocol that relies on proteinase K for cuticle digestion. The cuticle is a major hurdle for the isolation of molecules from worms and proteinase K provides an effective method to break it. However, proteinase K has to be heat-inactivated to be able to use enzymes for reverse transcription. Nanofluidic systems are ideal when investigating a given set of target transcripts and the use of either smaller (multi-array chip) or larger (single-array chip) number of samples allowing adaptation to the scale of the experiment. To obtain an unbiased picture of all transcripts expressed in a single worm, the obvious choice is to use RNA sequencing. If, however, the focus of the experiment is a smaller but still relatively large set of target genes, it is more cost-effective to utilize this protocol, provided the researcher has access to a nanofluidics PCR machine. The cost of the nanofluidic system reagents and a single-array chip is estimated as approximately £13 per worm, whereas the costs of the reagents for single-worm sequencing would be approximately £60 per worm, not including the sequencing costs.
When considering what PCR platform to use, the Worm-to-CT method coupled to nanofluidic qPCR offers advantages with regards to time and throughput. It is possible to obtain 9,216 RT-qPCR results in approximately 2 days of work, whereas amplification of the same number of targets using a standard qPCR platform would take approximately 5 working weeks using 96 well plate assays, running four plates a day. However, if the number of targets to be tested is smaller, then it is more cost-effective to use Worm-to-CT coupled with a standard qPCR machine. The single-array chips cannot be rerun, so running empty wells decreases cost-efficiency.
One limitation of the method is the potential formation of primer-primer dimers during the multiplexing step, but this occurs in less than 1% of the cases. Although the Worm-to-CT protocol is efficient and provides reliable results when applied to single worms, there is a failure rate of about 5%, which likely corresponds to cases where the worm remains trapped in the cap or the top of the tube during the harvesting step.
Together, this versatile and reliable method offers increased throughput and sensitivity compared to more standard techniques. This method can be very useful for validation of high-throughput screens and is an excellent choice to either monitor or validate single-worm gene expression levels. This method can be applied to other challenging techniques, such as the quantitation of gene expression from isolated tissues. For example, isolation of full tissues, such as the intestine, gonads, or cells isolated by FACs, provides enough material to perform RNA sequencing experiments. However, limited amounts of material often lead to duplicated reads, which precludes quantitation of rare transcripts. In this scenario, using nanofluidics-based technology should provide added sensitivity to the experiments and increase cost-efficiency if the researchers need to monitor only a subset of all transcripts in those tissues or cells.

Supplementary Material
Refer to Web version on PubMed Central for supplementary material. Comparison of Worm-to-CT protocol versus regular guanidium thiocyanate-phenolchloroform extraction 22 on bulk samples. Consistent with previous findings, in hsf-1(sy441) mutants 23 , the levels of hsp transcripts in response to heat shock decreased. The above histograms depict the induction of hsp-70 in the absence of (-), or following (+) a short heat shock of 30 min at 34 °C. The cDNA was obtained using guanidium thiocyanate-phenolchloroform extraction applied to 1,000 worms (left) or using the Worm-to-CT method applied to 30 pooled worms (right). The expression levels of hsp-70 per 100 ng of total RNA obtained by each method were compared. As expected, in hsf-1(sy441) the transcriptional induction of hsp-70 in response to heat shock significantly decreased by 82.7% using guanidium thiocyanate-phenol-chloroform and by 92.3% using the Worm-to-CT method. The mRNA levels from target genes were normalized against the average of the three housekeeping genes cdc-42, pmp-3, and ire-1. Each dot represents a biological replicate.   The data presented in this graph were obtained in 28 individual worms following a short heat shock (30 min at 34 °C). Each red dot represents the coefficient of variation (CV) of mean transcript expression levels for one transcript assayed between 28 individual worms (bio CV). Each blue dot represents the CV of expression levels between three technical replicates obtained from a single worm, per transcript assayed (technical CV). This graph shows that technical variability (between technical replicates) was much lower than biological variability (between individual worms), suggesting that it is unnecessary to perform technical replicates on a nanofluidic gene expression array when assaying gene expression in single worms, similarly to single-cell studies 14 Table 1 Plan layout for a multi-array chip.
The  Table 2 Plan layout for a single-array chip.
The table above shows a simple layout that can be utilized when planning a single-array chip run. On the left are spaces that should be filled with primer targets of interest and on the right are spaces that should be filled with the samples of interest.  Table 3 List of RT-qPCR primers used in this study.